1. health and fitness
2. therapy

640
Chem. Rev. 2011, 111, 640–656
How to Catch a Membrane Protein in Action: A Review of Functional
Membrane Protein Immobilization Strategies and Their Applications
Virginie Fru¨h,†,‡ Ad P. IJzerman,† and Gregg Siegal*,‡
Division of Medicinal Chemistry, Leiden Amsterdam Center for Drug Research, Leiden University, Leiden, The Netherlands, and Leiden Institute of
Chemistry, Leiden University, Leiden, The Netherlands
Received March 4, 2009
Contents
1. Introduction
2. Noncovalent Interactions with the Whole Cell
2.1. Via Wheat Germ Lectin-Agarose for
Whole-Cell ImmobilizationsMP: Glucose
Transporter Glut1, P2Y1, and H1R
3. Noncovalent Interactions (Adsorption) with the
Membrane
3.1. Via Adsorption to Polylysine-Ion Channel
5-HT3 Receptor and GPCR R1B-Adrenergic
Receptor
3.2. Via Direct Adsorption of Native Attoliter
Vesicles on Glass SlidessIon Channel: 5-HT3
Receptor
3.3. Via Native Membrane Hydrophobic Interactions
on BeadssMP: Acetylcholine Receptor
3.4. Via Electrostatic Immobilization of
ProteoliposomessMP: Glucose Transporter
Glut1
3.5. Via Direct Lipid Distribution on GlasssMP: fd
Coat and FXYD Proteins
3.6. Via Liposome Fusion into Lipid Bilayers on
Platinum/Glass or Silicon SlidessGPCR:
Bacteriorhodopsin
3.7. Via Microarrayed SurfacessGPCRs:
Adrenergic Receptor, Neurotensin Receptor,
and Dopamine Receptor
4. Covalent Interactions with Linker/Spacer to
Membrane
4.1. Via Thiopeptide Tethered Membrane
BilayerssMP: Cytochrome C Oxidase
4.2. Via Polymer Cushion-Supported Lipid
BilayerssGPCR: Rhodopsin
4.3. In Vitro Synthesis in the Presence of a
Tethered Lipid MembranesGPCR: OR5
4.4. Via Polymerized Lipid MonomerssGPCR:
Rhodopsin
5. Covalent Interactions with Protein with Subsequent
Lipid Reconstitution
5.1. Via Carbodiimide Coupling with Lipid
ReconstitutionsGPCR: Rhodopsin
6. Specific Immobilization of Protein in the Native
Membrane via Linker
640
642
642
642
642
645
645
646
646
647
647
648
648
648
649
649
649
649
650
* To whom correspondence should be addressed. Leiden Institute of
Chemistry, Leiden University, Postbus 9502, 2300RA Leiden, The Netherlands. Phone: +31 (0)71 527 4543. Fax: +31-(0)71-5274609. E-mail:
[email protected].
†
Leiden Amsterdam Center for Drug Research, Leiden University.
‡
Leiden Institute of Chemistry, Leiden University.
6.1. Via Biotinylated Protein Bound to Streptavidin
or AvidinsGPCR: Neurokinin-1 Receptor
7. Specific Immobilization of Detergent Reconstituted
Protein via Linker
7.1. Via N-terminus of Protein on Sepharose
ResinsMP: KcsA and DsbB
7.2. Via Tag and Biotinylated Antibody on
BSA-Biotin SurfacessGPCR: β2-Adrenergic
Receptor
7.3. Via HIS Tag on Quartz SurfacesMP: Ion
Channel 5-HT3R
7.4. Via Biotinylated LigandsGPCR: Neurotensin
Receptor-1
8. Specific Immobilization of Lipid Reconstituted
Protein via Linker
8.1. Via Biotinylated Proteins in a Mixed
Self-Assembled Monolayer (SAM)sGPCR:
Rhodopsin
8.2. Via Lipid Bilayer Tethered through HIS-Tagged
ProteinsMP: Cytochrome C Oxidase
9. Specific Immobilization of Alternatively
Reconstituted Protein via Linker
9.1. Via Biotinylated AmphipolsVarious MPs
9.2. Via the N-terminus of Nanodisc Solubilized
Protein MP: KcsA and DsbB
10. Perspectives
11. Abbreviations
12. Acknowledgments
13. References
650
650
650
651
651
652
652
652
653
653
653
653
654
654
655
655
1. Introduction
Analysis of the results from the human proteome project
suggests >30% of proteins are membrane bound.1 Membrane
proteins (MPs) are in the core group responsible for signal
transduction, including the G-protein coupled receptors
(GPCRs), which represent 30-40% of targets for marketed
drugs.2 It is needless to stress, therefore, the great commercial, industrial, and research value of MPs. There is
growing interest in immobilizing MPs to various surfaces
to create new biosensors or platforms enabling the study of
their biological functions and identification of new leads for
drug discovery. Soluble proteins have been readily immobilized through cross-linking aldehydes or thiols with
protein amines or carboxyl groups for applications such as
microarraying by printing.3-6 Functional immobilization of
MPs requires consideration of their physiological needs, often
dictated by the quality and components of the natural
hydrophobic environment surrounding this class of proteins.
10.1021/cr900088s  2011 American Chemical Society
Published on Web 09/10/2010
Functional Membrane Protein Immobilization Strategies
Virginie Fru¨h carried out her undergraduate studies in Switzerland and in
the U.K., where she graduated with a B.Sc. in coastal marine biology.
She went on to work for Serono Laboratories in Switzerland, contributing
to projects in both biochemistry and pharmacology. In 2009, she obtained
her Ph.D. in biochemistry at the University of Leiden under the supervision
of Dr. Ad. P. IJzerman and Dr. Gregg Siegal. She is currently a
postdoctoral fellow at the Agricultural University of Bogor, Indonesia,
working on the development of sustainable surfactants.
Ad IJzerman is professor of medicinal chemistry at the Leiden/Amsterdam
Center for Drug Research (Leiden University) and an expert on G proteincoupled receptors, in particular adenosine receptors. His team, together
with the Stevens’ lab at the Scripps Institute, has recently achieved the
structure elucidation of one of the adenosine receptor subtypes, the A2A
receptor. He has a keen interest in novel concepts of intervening with
receptor function, in particular allosteric modulation and residence time.
The aim of this review is to focus on the immobilization
chemistry applied to MPs and how these have enabled ever
more complex assays from simple whole cell analyses to
purified proteins on a chip, from the 1990s onward.
With MPs, often the challenge is not the immobilization
per se, but rather the requirements of the analysis and the
protein itself with regards to protein availability and the type
of lipid environment needed. Earlier studies of MPs, because
of the difficulty in purifying them, involved whole cell or
membrane fragment immobilization where native biological
responses could be followed by “macroscopic” methods such
as frontal chromatography (section 2). In these studies,
immobilization was accomplished by simple techniques such
as incorporation of membrane fragments upon swelling of
gel beads during freeze-thawing7,8 or by coating beads with
various cell-adhesive substances.9 Although these systems
allow one to study the native environment, the immobilization itself is random and relatively unstable. Furthermore,
reliable signal detection is often limited by high levels of
nonspecific binding to the environment around the target MP.
Chemical Reviews, 2011, Vol. 111, No. 2 641
Dr. Siegal obtained his Ph.D. at the University of Rochester in New York
and pursued postdoctoral studies with Professor Kurt Wu¨thrich at the ETH
in Zu¨rich, Switzerland, and Professor Paul Driscoll at the Ludwig Institute
of Cancer Research in London, U.K. In 1997, he moved to Leiden
University, where he received a Dutch Royal Society Fellowship to form
his own research group. His current research interests include development
of drug discovery technologies and their implementation in the drug
discovery process. Recent efforts have focussed on application of
biophysical ligand screening techniques to membrane proteins.
Solubilized MPs may be either used as such in detergent
micelles or reconstituted in various types of detergent or lipid
formations.10-13 With recent advances in the development of
mimics of the native membrane, however, the stability and
formation of various lipid assemblies are becoming less
problematic. However, one cannot emphasize enough how
challenging these steps are. Removing the protein from its
hydrophobic environment and reconstituting it into another is
a difficult process that may damage the protein or result in less
than optimal functionality.14,15 Furthermore, increasing complexity arises with increasing numbers of transmembrane
segments.16 The most successful stories are currently based
on the nicotinic acetylcholine receptor and rhodopsin, due
to the availability of these MPs in large quantities and
relatively good stabilities. Newer membrane mimics have
allowed direct adsorption of appropriately prepared MPs to
flat surfaces or chips made of glass, platinum-glass, and
platinum-silicon to create monolayers of lipids that hold
the protein onto the surface (section 2). Subsequent strategies
have evolved to create even more stable lipid bilayers, as
opposed to less stable monolayers, by fusing various types
of solubilized MPs to lipid monolayers on surfaces. These
methods provide more robust immobilization and are applicable to more sensitive studies, provided fluorescently or
radioactively labeled components are available to specifically
monitor MP activity. As a result, the highly sensitive
detection of such signals does not require large amounts of
solubilized MP. As these methods were applied to increasing
numbers of MPs, it became clear that immobilizing membranes or mimics too close to a surface may restrict diffusion
of solutes below the membrane and sterically hinder extramembranous protein dynamics. As a result, so-called solidsupported membranes have been developed, where the space
below has been filled with hydrophilic polymers, combined
with various linkers, to immobilize the MP in a manner that
accommodates dynamic behavior. Solid-supported membranes are a better mimic of the natural environment because
they create additional membrane fluidity and allow for
mobility of large extramembranous protein domains17 (section 3). Many of these techniques have evolved in such a
way that solubilization and purification of the protein may
not be a prerequisite, but when this is the case, the
642 Chemical Reviews, 2011, Vol. 111, No. 2
solubilization from crude cell lysates or lipid reconstitution
may be carried out in situ (section 4).
On the detection side, earlier assays primarily used
biological readouts. Although these techniques were sufficient for high-throughput or high-content assays of, for
example, ligand binding and cell signaling activation, they
typically required labeling of a protein or ligand to produce
a signal. Recently, newer, so-called label-free technologies
have emerged, such as surface plasmon resonance (SPR) and
NMR,18 that are able to directly detect ligand binding, thereby
rendering use of chemically, biologically, or radioactively
labeled protein or ligands redundant. These technologies are
sensitive to intermolecular interactions and capable of
detecting weak binding affinities in the mM range (for NMR),
provided the target protein is available in sufficient quantities
and functionally solubilized. As opposed to soluble proteins,
MP interactions with ligands, lipids, or other proteins are
often specifically located on one side of the cellular
membrane. Some strategies have therefore been taken one
step further by using high-affinity interactions for specifically
orienting MPs in the immobilized membrane on chips
(sections 5-7). Techniques involving in vitro expression in
the presence of the tethered lipid membrane in which the
protein will be integrated directly upon expression19 are an
elegant way to achieve oriented, immobilized MPs.
As strategies to couple MPs become increasingly perceptive, a thorough summary of the numerous accomplishments
is useful. Here, we present such a summary in the form of
text and tables (see Tables 1 and 2) that focuses on the
chemistries of various immobilization types, with descriptions
of their applications and results.
2. Noncovalent Interactions with the Whole Cell
2.1. Via Wheat Germ Lectin-Agarose for
Whole-Cell ImmobilizationsMP: Glucose
Transporter Glut1, P2Y1, and H1R
Fru¨h et al.
The Kd of CB and D-glucose binding to Glut1 were comparable
to values previously reported from studies on whole cells and
membrane vesicles. This immobilization method also enabled
comparison of the Kd of Glut1 for CB and D-glucose across
different types of preparations of the reconstituted receptor,
without any effects on the affinity. The advantage of this
technique was that cells remained intact, in contrast to electrostatic adsorption, which causes hemolysis.15 The gel beads
could be stored for one month but lost 40% of their capacity
to bind cells. It was reported that it was not cell integrity
but rather the capacity of cells to bind to the gel (and
therefore the remaining functionality and density of membrane glycophorins) that determined the stability of the
column, along with physical and chemical properties of the
storage environment. Another advantage was that protein
purification was not necessary, and manipulations involving
column preparation and cell immobilization were simple.
Nevertheless, the immobilization was sensitive to pressure
changes, and gentle washing induced detachment of the cells,
causing loss in reproducibility. Using this immobilization
strategy with other applications will depend on conditions
for cell homeostasis and functionality, as this technique was
only optimized with respect to hematocyte cell lines. A
variant of the whole cell immobilization technique involves
immobilization of cytoplasmic membrane preparations on
silica beads that have been derivatized with lipids. This socalled cellular membrane affinity chromatography has been
used to noncovalently immobilize membrane suspensions
from mammalian cells recombinantly expressing GPCRs. In
particular, membrane preparations from a human astrocytoma
cell line, 1321N1, expressing the P2Y1 purinergic receptor,
wereimmobilizedonimmobilizedartificialmembrane-phosphotidylcholine (IAM-PC) columns.21 The authors first demonstrated specific binding of 2-MeSADP to the immobilized
P2Y1 receptor and determined a Kd that compared well to
literature values. Subsequently, they probed for the presence
of endogenously expressed histamine H1 receptor by assessing the binding of Mepyramine. Mepyramine bound with
a Kd of 2.9 nM, in good agreement with literature values,
demonstrated that multiple GPCRs could be assayed by
immobilization of membrane preps from a single cell line.
The columns of immobilized membranes proved to be stable
for 6-8 weeks.
3. Noncovalent Interactions (Adsorption) with the
Membrane
Frontal chromatography can be used to study interactions
between a mobile phase and a stationary phase by regression
analysis of the retention volumes. Furthermore, when whole
cells are immobilized, this technique can allow the study of
binding affinities of soluble ligands to MPs in their native
environment. Immobilization can be achieved by adhering
whole cells to agarose gel beads via wheat germ lectin
(WGL).9,15 WGL is a derivative of wheat germ agglutinin and
binds to N-acetylglucosamine and to N-acetylneuraminic acid
on the extracellular face of red blood cell membranes.20 The
Glut1 transporter targeted for these assays was indirectly
immobilized to Sepharose 4B gel beads prederivatized with
WGL by simply incubating cells overexpressing the receptor
with gel beads.20 The interaction of Glut1 with D-glucose
and cytochalasin B (CB, a fungal alkaloid that inhibits
glucose transport) was measured by frontal chromatography.
3.1. Via Adsorption to Polylysine-Ion Channel
5-HT3 Receptor and GPCR r1B-Adrenergic
Receptor
Studying whole cells by fluorescence often leads to high
background signals from the cytoplasm. This issue, combined
25
30
36, 37
1
43-45
46, 48
13, 56
61, 63
19
64
10
12, 73-75
79
12
88
89
93
94, 95
102
80
3.2
3.3
3.4
3.5
3.6
3.7
4.1
4.2
4.3
4.4
5
6
7.1
7.2
7.3
7.4
8.1
8.2
9.1
9.2
ref
9, 15
22-24
2.1
3.1
section
R
R
O
O
O
O
O
O
O
D
R
D
R
R
R
R
O
I
R
R
I
I
I/R
covalent coupling
affinity capture
affinity capture
microprinting/affinity
capture
affinity capture
affinity capture
affinity capture
covalent coupling
in vitro expression/bilayer
insertion
cross-linking of bis-SorbPC
by UV light
microprinting/affinity
capture
affinity capture
hydrophobic adsorbance
hydrophobic tethers
hydrophobic adsorption
by printing
freeze-thawing and hydrophobic
glass adsorption
hydrophobic, liposome adsorption
onto monolayers
freeze-thawing
lectin-gel bead adsorption
polylysine, silicon nitride
interactions with native
membrane
native vesicle budding
from whole cells
hydrophobic adsorption
method
silver modified to bind
nickel (NTA)
biacore chip and
polysterene beads
sepharose resin with
reactive aldehyde groups
gold sensor chips
streptavidin-coated chip
glass modified to
bind nickel (NTA)
carboxymethyl-modified
dextran
glass slides chemically
oxidized silicon
sepharose resin with reactive
aldehyde groups
glass or gold surfaces
cuvette
gold-covered chromium
layered glass
carboxymethyl-modified dextran
polymer hydrogel
gold
γ-aminopropylsilane
(GAPS)-derivatized gold slides
platinum/glass and
platinum/silicon slides
glass slides
glass slides and/or
functionalized glass
polyvinyltoluene fluorophor
microbeads
superdex 200 gel beads
Sepharose 4B gel beads
polylysine coated glass or
silicon nitride surfaces
surface
5-carbon chainshydrophilic
streptavidin-biotin
protein HIS tag
thiolipid and biotin
streptavidin-biotin
streptavidin-avidin +
antibody + flag tag
protein HIS tag
streptavidin-avidin +
antibody
5-carbon chainshydrophilic
peptides
polymer cushion
thiopeptides
tethers
synthetic polymer
Amphipol (Apol)
nanodiscs
nonaethyleneglycol
monododecylether (C12E9)
micelles
CHAPS/DDM/CHS
detergent
self-assembled monolayers
(SAMs) of w-hydroxyundecanethiol
(HTA)
reconstituted DPGPC
DDM detergent
DPC detergent
reconstituted POPC
lipid bilayer
native membrane vesicles
covalent peptidesDMPE
fused with PC vesicles
OG vesicles
Triton X-114, Triton X-100,
sodium cholate
native membrane vesicles
native vesicles or
proteoliposomes
reconstituted
POPC/POPG
synthetic bilayers formed
by fusion of proteoliposomes
to monolayers
vesicles of DPPC/DMPC
(4:1) egg-yolk PC
native membrane vesicles
native membrane
native membrane
native membrane
membrane
OmpA KcsA DsbB
bacteriorhodopsin
cytochrome C oxidase
rhodopsin
Neurotensin receptor-1
β2 adrenergic receptor with
FLAG tag and fluorophore
5-HT3R
biotinylated neurokinin-1
receptor, Gramicidin-A
OmpA KcsA DsbB
rhodopsin
rhodopsin
OR5
rhodopsin
rhodopsin, acetylcholinesterase,
cytochrome oxidase, nicotinic
acetylcholine receptor
adrenergic receptor,
Neurotensin receptor,
dopamine receptor
cytochrome C oxidase
Fd coat protein
glucose transporter Glut1
acetylcholine receptor
5-HT3 receptor
glucose transporter Glut1
5-HT3 receptor
protein
Table 1. Summary of Direct (D) or Indirect (I) Immobilization Methods with Random (R) or Oriented (O) Protein on the Surface, Including Type of Membrane and Protein Involved, In
Order of Appearance in the Text
Functional Membrane Protein Immobilization Strategies
Chemical Reviews, 2011, Vol. 111, No. 2 643
22-24
25
30
36, 37
1
43-45
46, 48
13, 56
61, 63
19
64
10
12, 73-75
79
12
88
89
93
94, 95
102
80
3.1
3.2
3.3
3.4
3.5
3.6
3.7
4.1
4.2
4.3
4.4
5
6
7.1
7.2
7.3
7.4
8.1
8.2
9.1
9.2
ref
9, 15
2.1
section
cytochrome C oxidase
bacteriorhodopsin, OmpA,
cytochrome b6f, bc1,
NAChR
OmpA, KcsA, DsbB
Neurotensin receptor-1
rhodopsin
β2 adrenergic receptor with
FLAG tag and fluorophore
ion channel 5-HT3R receptor
rhodopsin
biotinylated neurokinin-1
receptor
OmpA, KcsA, DsbB
rhodopsin
OR5
rhodopsin
cytochrome C oxidase
rhodopsin,
acetylcholinesterase,
cytochrome oxidase,
nicotinic acetylcholine
receptor (nAChR)
adrenergic receptor,
Neurotensin receptor,
dopamine receptor
Fd coat protein
glucose transporter Glut1
acetylcholine receptor
ion channel 5-HT3 receptor
NK1 receptor and ion channel
5-HT3 receptor
glucose transporter Glut1
protein
2
100 µmol/mL
250 RU
100 nmol/mL
0.1 pmol/mm
100-400 receptors/um2
400 RU
50 ng/cm2
12.5-125 ng/mg
1750 receptors/µm2
44.6 µg/mL
surface covered
solubilized
DPC
DDM
DDM, OTG, NMP, C8E4
nonaethylene glycol
monododecyl ether
detergent (C12E9)
CHAPS/DDM/CHS
no
DDM
DPC
egg PC
no
no
no
Triton X-114, Triton X-100,
sodium cholate, and
saccharose
no
no
OG, DPPA, DPPE, DPPC,
and cholesterol
cholate
no
no
no
no
no
one month
several hours
one month
few months
few months
several weeks
one month
shelf life
fragment-based drug
discovery, binding assays
binding
binding studies, activation
with agonist
redox experiments
binding experiments
fragment-based drug
discovery
binding studies, activation
under agonist
binding
potential for hybridization,
kinetics, and binding
binding studies, protein
orientation
isomerization and G-protein
activation
binding studies
binding studies
deorphanizing receptors, lead
optimization and
identification, bioassays,
high-throughput screening
binding studies and kinetics
binding interactions,
interactions with other
components in the cell
membrane
binding studies, identification
of orphan receptors
binding, interactions,
conditions
structural solid-state, 15N
labeled protein
binding studies
binding, interactions,
conditions
lipid and protein mobility in
the native membrane,
binding Interactions
studies allowed
NMR, fluorescence-based
assays
electrochemistry
surface plasmon resonance
SPR
SPA
total internal reflection
fluorescence
plasmon-waveguide resonance
(PWR) spectroscopy
SPR
total internal reflection
fluorescence
NMR and fluorescence-based
assays
fluorescence microscopy
SPR, impedance
spectroscopy, AFM
SEIRAS
impedance spectroscopy
fluorescence-based assays,
radioligand binding assays*
radioligand binding,
electrochemistry,
fluorescence
solid-state NMR
frontal chromatography
fluorescence, fluorescence
after photobleaching
(FRAP), single-molecule
imaging
fluorescence, confocal
microscopy, radioligand
binding, fluorescence
imaging
radioligand binding
frontal chromatography
analysis
Table 2. Summary of Protein Immobilization Solubilization Requirements, Shelf Life, Type of Analysis Obtained, And Type of Analytical Methods Applicable, In Order of Appearance in the
Text
644 Chemical Reviews, 2011, Vol. 111, No. 2
Fru¨h et al.
Functional Membrane Protein Immobilization Strategies
with limited access to the intracellular compartment, obscures
many of the dynamic processes between the membrane
leaflets. To circumvent this, supported cell-membrane sheets
allow one to observe target proteins from the intracellular
side when both leaflets of a cell membrane are physically
pulled apart.22,23 The ligand-gated ion channel 5-HT3 receptor
and the R1B-adrenergic receptor were both overexpressed in
HEK-293 cells, which were grown on poly-L-lysine (PLL)coated plastic. Another PLL surface was pressed down onto
the cells and removed after several minutes of contact,
ripping apart the cells and exposing the membrane’s intracellular face with endogenous lipids and proteins. To
visualize the target lipids and proteins, the latter were
expressed as fusion constructs with fluorescent proteins and
the former were colocalized with fluorescent lipid markers.
The supported polylysine membrane sheets have also been
modified to allow access to the extracellular side by pressing
down silicon nitride surfaces into which arrays have been
cut out.24 Labeling with fluorescent probes from different
sides of the membranes enabled the authors to study the
mobility of lipids, GPCRs, and G proteins within and
between the leaflets.23 Following the dynamics of heterotrimeric G protein partitioning into lipid anchor microdomains
was also possible along with the mobility regimes of
glycosylated transmembrane proteins. Competition binding
studies using known, fluorescent ligands confirmed the
functionality of the proteins studied. In comparison to wholecell analyses, the advantages of supported cell-membrane
sheets are numerous. The mechanical stability is excellent,
minimizing unwanted movement of the observed membrane
upon application of stimuli, which can be problematic for,
e.g., imaging applications. Although labeling is required,
specific targets can be observed in their native surroundings,
even maintaining the appropriate orientation, while all the
time monitoring cellular processes such as signaling and
protein translocation. Following populations of labeled
proteins was therefore possible as a direct result of removing
the intrinsically highly fluorescent cytosol during the preparation of these membrane sheets. This strategy is generally
applicable to all cell membranes but is restricted to the use
of labeled ligands or proteins. Adapting the immobilization
to metallic surfaces may allow the strategy to be applied to
surface plasmon resonance applications.
Chemical Reviews, 2011, Vol. 111, No. 2 645
or via biotinylation of membrane components or target
proteins.26,27 Attoliter vesicles can be obtained in one step,
without changes in receptor function, orientation, or localization in the membrane. Vesicle formation is a simple process
induced by incubating cells, in this case expressing the
serotonin-gated mouse ion channel 5-hydroxytryptamine
type-3 receptor (5-HT3R), with cytochalasin B (CB). This
causes cells to produce tubular extensions that bud off when
agitated. Native vesicles were thus formed that could be
separated from whole cells by centrifugation. Usefully, the
vesicles could be stored at 20 °C for weeks without causing
loss of receptor function. Coexpression of cytosolic green
fluorescent protein resulted in the possibility to monitor and
characterize vesicle formation and the presence of cytosolic
components. Specific binding of fluorescently labeled ligands
could be used to locate the receptor in the vesicles, and
radioligand binding assays revealed their appropriate orientation and function. Furthermore, agonist and antagonist
ligands showed the same behavior with vesicles as with
detergent-solubilized receptors. Finally, Ca2+ signaling within
the vesicles upon agonist activation could also be detected,
suggesting the functionality of the complete signaling
pathway. These micrometer-sized vesicles are novel miniaturized reaction centers where receptor-based assays can be
carried out in physiological conditions. These experiments
not only revealed the native function of the receptors within
these vesicles but also proved that other important players
such as ion pumps are still present and functional. Vesicle
formation yielded native behavior when tested with Chinese
Hamster Ovary (CHO) or HEK293 cells, suggesting this
procedure can be generally applied to other cell strains. When
fluorescent ligands are available, these miniaturized and
stabilized compartments can provide the basis for highthroughput assays where patterns of immobilization can
reveal receptor function and binding affinities depending on
various tailored reactive groups.
3.3. Via Native Membrane Hydrophobic
Interactions on BeadssMP: Acetylcholine
Receptor
3.2. Via Direct Adsorption of Native Attoliter
Vesicles on Glass SlidessIon Channel: 5-HT3
Receptor
Obtaining large amounts of MPs is often limiting, and
therefore, immobilizing miniaturized vesicles containing the
target protein can be useful for studying receptor-mediated
cellular responses. In this particular case, Pick et al. have
been able to produce vesicles straight from the native
membrane of mammalian cells, with unprecedented attoliter
volumes.25 The vesicles can be immobilized in different
manners, either through direct lipid adsorption to glass slides
Radioligand binding assays are often used to quantify
binding affinities of ligands to MPs because only picomolar
amounts of protein are required. However, this requires
separation of bound from unbound ligands. To circumvent
this, the scintillation proximity assay system (SPA)28,29 was
developed, where receptors are immobilized on solid microspheres that contain scintillation fluid. This homogeneous
assay provides specific information on radiolabeled ligand
binding without the need for separation from unbound ligand,
as scintillation is only stimulated by radioligand in close
proximity to the microspheres.30,31 As a first attempt to apply
SPA to MPs, membranes containing acetylcholine receptor
were isolated from the electric organ of Torpedo californica
646 Chemical Reviews, 2011, Vol. 111, No. 2
and directly added to polyvinyltoluene fluorophore microbeads previously suspended in Triton X-100. Under these
conditions, the membrane preparations spontaneously adsorbed to the beads. These beads contained scintillation fluid,
and upon binding of the radiolabeled ligand to the receptor,
the close proximity of receptor led to photon emission
proportional to the amount of immobilized protein. In
addition to obviating the requirement for separation of
unbound from bound ligand, this method has less nonspecific
binding than usually encountered in radioligand binding
assays. Radioligand binding assays of 125I-R-bungarotoxin
to the acetylcholine receptor enabled the study of various
parameters such as dose response of acetylcholine and
suberyldicholine with respect to radiolabeled R-bungarotoxin
binding, providing apparent Ki values. The effects of different
parameters such as concentrations of detergents and salts on
the activity of the receptor were also studied, along with the
detection of specific antigens by antibody binding. SPA has
a number of other advantages for MPs. Because of the great
sensitivity of scintillation counting, as little as 1 ng of
receptor can be detected. Although SPA requires prior
radiolabeling and modification of the protein or ligand, many
receptor-SPA beads are available commercially. The method
has been used in a broad range of studies, varying from
identification and quantification of PCR products32 to highthroughput screening of inhibitors of DNA binding proteins33,34
to studying interactions between antibodies and antigens.35
Finally, because of the simplicity of the immobilization
procedure, it should be generally applicable to all MPs.
Fru¨h et al.
kinetics were easily studied with such a setup due to lower
nonspecific binding to the lipids in vesicles compared to
whole cells. Immobilized membrane vesicles had 4 times
more transport inhibitor (CB) binding capacity than proteoliposome gel beads, showing that the membrane vesicles
contained a high density of Glut1. In the presence of
dithioerythritol (DTE), the binding capacity of CB was >80%
stable after 20-40 days in all types of columns studied.
Furthermore, the activity could be controlled by inhibiting
Glut1 with HgCl2 and restoring Glut1 activity by adding
DTE, which has the effect of partially reversing the Hg2+
inactivation. Steric trapping should enable other applications
such as studies of effects of ionic strength, pH, temperature,
and lipid composition on ligand-binding characteristics. This
methodology has the potential, as do other whole cell or
membrane fragment-based applications, of enabling studies
of interactions occurring within the native membrane because
of the presence of other required membrane-bound components. However, applications of this immobilization strategy
remain limited to the use of gel beads for steric trapping of
the swelled vesicles, where high amounts of protein per mg
of membrane are required for sensitive readouts. Furthermore, the noncovalent immobilization leads to degradation
during use, and clearly, the protein to be studied needs to
be stable to multiple freeze-thaw cycles.
3.5. Via Direct Lipid Distribution on GlasssMP:
fd Coat and FXYD Proteins
3.4. Via Electrostatic Immobilization of
ProteoliposomessMP: Glucose Transporter Glut1
Quantitative affinity chromatography used to be limited
to studying interactions between soluble substances. By
immobilizing a membrane protein on gel beads, however,
binding of soluble substances to MPs could be quantified
by regression analysis of the retention volumes for different
concentrations of ligands. To study interactions between
soluble substances and the glucose transporter Glut1, membrane vesicles containing the latter were immobilized onto
gel beads.36,37 Membrane vesicles were obtained by stripping
red blood cells of peripheral proteins and partially solubilizing them with octylglucoside (OG). In some cases, the
vesicles were purified by ion-exchange chromatography for
reconstitution of the Glut1 transporter into proteoliposomes.38
It has been reported that freeze-thawing39 and freeze-drying
with subsequent rehydration40 can cause vesicles or proteoliposomes to fuse together and form larger particles. This
approach was used to sterically trap vesicles after two cycles
of freezing in ethanol/CO2 and thawing in the presence of
gel beads.36 Using the sterically trapped liposomes, specific
interactions between Glut1 and CB, and between Glut1 and
D-glucose,41 were observed. Ligand binding affinities and
The biological role of the protein coating filamentous fd
bacteriophages (fd coat protein) involves interacting with the
host’s cellular membrane. Three-dimensional structure determination is a powerful method to determine molecular
mechanisms of protein function. However, in many cases,
solution NMR studies of an MP in detergent micelles will
not provide sufficient information because the dynamics and
orientation of the protein in detergent micelles may be very
different than in a membrane bilayer. Therefore, the fd coat
protein was immobilized onto glass slides in phospholipid
bilayers with one simple step for structural studies using
solid-state NMR.1 For these studies, 15N-labeled fd coat
protein was produced in E. coli and solubilized in detergent
micelles. The solubilized fd coat protein was then reconstituted into phospholipid vesicles consisting of a mixture of
palmitoyl oleoyl phosphatidylcholine/palmitoyl oleoyl phosphatidylglycerol (POPC/POPG).42 Glass slides were covered
with the vesicle suspension containing the reconstituted
protein, and the bulk water was left to evaporate. The slides
were then stacked and bathed in saturated ammonium
phosphate solution. Finally, the slides were placed in thin
polyethylene tubing and inserted into a rotor for NMR
studies, which allowed complete resolution of the amine
resonances in a 3D magic angle spinning (MAS) NMR
correlation spectrum. The choice of lipids determines the
Functional Membrane Protein Immobilization Strategies
level of bilayer orientation on the slides and allows control
over the horizontal and vertical spacing between phospholipids.11 The presence of unsaturated chains leads to more
fluid bilayers, and larger vertical spaces between bilayers
can be obtained by incorporating negatively charged lipids.11
The noncovalent, hydrophobic interactions with the glass
were strong enough to hold the vesicles immobile on the
substrate for the duration of an NMR experiment, despite
the considerable shear forces generated by spinning the
sample. No details were given on the protein stability or
functionality, but it was fully incorporated into phospholipid
bilayers onto the glass support. This immobilization strategy
allows a wider range of MPs to be studied by solid-state
NMR and may be used in other applications compatible with
glass surfaces. The extensive vesicle preparation was a
complicated process, however, and may not be generally
applicable to other MPs. Although fd coat protein could be
obtained in relatively large quantities, this is rarely the case
with other MPs. Further, it may prove difficult to adapt this
method of orienting the protein to new, small-volume MAS
rotors.
3.6. Via Liposome Fusion into Lipid Bilayers on
Platinum/Glass or Silicon SlidessGPCR:
Bacteriorhodopsin
For an immobilized MP to remain functional, the fluid
lipid or detergent environment must not become too rigid
upon immobilization or it might impede the dynamic
behavior of MPs. Lipid rigidification can be minimized when
proteoliposomes are immobilized by fusion with monolayers
on glass surfaces to form bilayers without any chemical
cross-linking. To demonstrate the approach, bacteriorhodopsin from purple membranes of Halobacterium halobium was
added to a lipid, cholesterol, and octylglucoside mixture, and
upon removal of octylglucoside during gel filtration, proteoliposomes were formed by the so-called detergentdepletion technique.43 Langmuir-Blodgett (LB) lipid monolayers44 were used to treat platinum/glass surfaces and
subsequently converted into bilayers when fused with the
bacteriorhodopsin proteoliposomes.45 Bacteriorhodopsin photoactivity in the reconstituted bilayers, monitored by electrochemistry, was comparable to natural membrane fragments
containing bacteriorhodpsin on platinum surfaces. Three
other MPs were also studied, including acetylcholinesterase
from bovine brain, cytochrome oxidase from bovine heart,
and the nicotinic acetylcholine receptor from the electric
organ of Torpedo. The purification and incorporation into
proteoliposomes of the three MPs varied depending on
suitable detergents, but they were all incorporated into stable
lipid bilayers following the same protocol. Acetylcholinesterase enzymatic activity was easily studied by fluorescence
measurements despite low immobilization yields, but only
semiquantitative data was available for cytochrome C oxidase, whereas studies of the nicotinic acetylcholine receptor
Chemical Reviews, 2011, Vol. 111, No. 2 647
with its ligand 125I-R-bungarotoxin were hindered by issues
of nonspecific binding. Although this technique provided a
surface with a fluid membrane environment, meeting criteria
for efficient molecular assays and potentially providing access
to printing arrays, the usual challenge of obtaining functional
protein in proteoliposomes remains. The bilayers did remain
immobilized to the supports for prolonged periods of time
after storage, but no information was provided on the stability
of the immobilized MPs.
3.7. Via Microarrayed SurfacessGPCRs:
Adrenergic Receptor, Neurotensin Receptor, and
Dopamine Receptor
Reducing the demand for protein and reagents has been
one of the strategies used to address the limitations faced
when MP purification and solubilization result in low
yields. This limitation is even more critical when MP
immobilization is at the heart of high-throughput screening
for drug discovery. An innovative way of addressing both
limitations has been to immobilize GPCRs through microarray printing assays.46,47 A nice example of this
technique has been developed at Corning. Membranes in
vesicular solutions of dimyristylphoshatidylcholine (DMPC)
and 1,2-dimyristoyl-sn-glycerophosphatidylethanolamine
(DMPE) or egg-yolk PC were printed via a robotic-pin
printer on γ-aminopropylsilane (GAPS)-derivatized gold
surfaces.48,49 The details of the methods used with GAPS
are proprietary to Corning. GAPS-coated slides exhibited
high mechanical stability during printing, with no dependence on lipid phases, thereby overcoming the desorption
encountered when withdrawing the slides through air.50
While covalently attaching membranes has been noted to
impede lateral fluidity of membrane lipids, and consequently
protein function, noncovalent printing techniques, through
membrane adsorption onto glass slides, have proved to
preserve GPCR and G protein functionality. Thus, binding
constants of known ligands were readily obtained, as well
as compound selectivity between and within GPCR families.
However, the assay in this format could not be used to
differentiate between agonists and antagonists. Arrays of the
β1-adrenergic receptor, the Neurotensin receptor (NTR1), and
the dopamine (D1) receptor have been created. As an
example, upon incubation with fluorescently labeled neurotensin, fluorescence was restricted to areas in which NTR1
had been immobilized, as expected. Binding constants
consistent with literature were also determined for known
small-molecule ligands using the printed array.48,49 A recent
exciting advance in printing is the adaptation to porous
substrates onto which the membranes are transferred, allowing access to both sides of the membrane.51 Array printing
is expensive because of the costs involved in acquiring such
printers and also in time spent to optimize procedures that
involve correct buffer compatibility, ligand specificities,
648 Chemical Reviews, 2011, Vol. 111, No. 2
affinities, and association and dissociation rates. However,
although at present microarrays have been analyzed with
fluorescent or radiolabeled46 ligands, they can in principle
be adapted for other applications such as SPR and electrochemistry, providing the surfaces are compatible. Information
on cell-surface interactions and ligands for orphan receptors
might also be derived from these microarrays, potentially
enabling discovery and validation of new lead molecules and
high-throughput screening. One interesting variant of this
idea is to generate a self-sorting array. Bailey and colleagues
printed arrays in which oligonucleotides of differing sequence
were localized onto appropriately prepared surfaces.52 Liposomes were generated from Sf9 cells recombinantly expressing either the H1 histamine or M2 muscarinic receptor.
Subsequently, cholesterol conjugated to an oligonucleotide
complementary to one of the immobilized oligonucleotides
was allowed to bind the liposomes. Incubation of a mixture
of liposomes with the printed surface resulted in sequencedependent immobilization of the GPCRs at the expected sites.
The GPCRs immobilized in such a manner bound known
ligands with expected affinities.
4. Covalent Interactions with Linker/Spacer to
Membrane
Fru¨h et al.
been established before, but it was measured here for the
first time using impedance spectroscopy. This immobilization
strategy proved to be compatible with sensitive measurement
of proton transport through the lipid bilayer. Furthermore,
kinetics and binding interactions of cyanide with CcO could
be studied. Because of the reported difficulty in preparing
thiopeptide or thiolipid compounds in the bulk phase, this
strategy allowed the simplified formation of a robust tethered
bilayer in steps. Furthermore, the tethered bilayer allowed
surface analytical techniques involving electric currents due
to the metal surface used as a support, suggesting the
technique may be adaptable to other applications with such
requirements. Again, the challenge is to find the appropriate
solubilization condition for the protein, but the space
conferred between the surface and the bilayer environment
of the protein by the tethered membrane bilayer can minimize
steric hindrance of large, dynamic MPs and allow smallmolecule transport and diffusion.
4.2. Via Polymer Cushion-Supported Lipid
BilayerssGPCR: Rhodopsin
4.1. Via Thiopeptide Tethered Membrane
BilayerssMP: Cytochrome C Oxidase
SPR applications require flat, metallic surfaces upon which
proteins must be immobilized at very high densities and,
ideally, as close to the surface as possible. However, studies
of immobilized MPs have revealed the need to create space
between the protein and the surface to reduce steric hindrance
of MP dynamics. Therefore, molecules such as thiolipids53
and thiopeptides have been developed as linkers, also called
tethers, because they couple the membrane in close proximity
to the surface but allow the membrane to remain fluid.
Thiopeptides have been used previously54,55 as a more
hydrophilic alternative to thiolipids, for example, to immobilize cytochrome C oxidase (CcO).13,56 In this particular
case, a thiopeptide-lipid monolayer was covalently linked
to gold surfaces with subsequent incorporation of the MP
within the lipid bilayer. An oriented peptide monolayer was
created by covalently binding an N-terminally thiolated,
oligoserine peptide to a gold surface.57 To complete the
tethered membrane mimic, vesicles obtained by dialyzing
Lipoid Egg PC58,59 were incubated with the monolayer
surfaces to form the thiopeptide-lipid bilayer by adsorption.
CcO, previously solubilized with Triton,56,60 was added to
the cell containing the thiopeptide-lipid bilayer surface and
spontaneously became incorporated into the bilayer with the
micellar dilution technique. All steps of the process could
be monitored by SPR, and final thicknesses of bilayers could
be measured. Active proton transport driven by CcO had
As discussed, previous studies revealed how immobilizing lipid bilayers containing MPs too close to the surface
may hinder protein dynamics and membrane stability. An
alternate solution to tethered membrane bilayers has been
the introduction of a hydrophilic polymer cushion interspaced with hydrophobic anchors below the lipid bilayer.
The cushion, formed by a hydrophilic polymer interspaced
with long alkyl chains, provides a lubricating surface that
allows the lipid bilayer to remain mobile and provides a
better mimic of the native environment.61,62 The gold L1
SPR chip used for such polymer-supported lipid layers
consisted of a covalently linked, carboxymethyl-modified,
dextran polymer hydrogel where a large part of the sugar
moieties were grafted with lipophilic alkyl side chains to
which liposomes adhere.61,63 Whereas the preparation of
small unilamellar vesicles (SUVs) containing rhodopsin
used to be necessary,63 here, fusion of crude rhodopsinenriched membranes directly onto Biacore L1 chips was used
instead of purification.61,63 SPR, atomic force microscopy
(AFM), and electrochemical impedance spectroscopy (EIS)
were used to follow receptor immobilization. The receptors
remained mobile in the plane of the lipid, suggesting that
this strategy has the potential to be used to study receptor
dimerization or interaction with other proteins.63 Furthermore,
the chips could be used directly, without prior treatment, and
repeatedly with different receptors after washes with detergents to remove earlier immobilized proteins. This may be
generally applicable to other systems as the strategy allows
on-surface enrichment for MPs with low expression yields.
Functional Membrane Protein Immobilization Strategies
Chemical Reviews, 2011, Vol. 111, No. 2 649
4.3. In Vitro Synthesis in the Presence of a
Tethered Lipid MembranesGPCR: OR5
Biophysical analysis requires pure protein preparations
functionally immobilized to a surface, and obtaining such
conditions for GPCRs remains challenging due to loss of
functionality and refolding problems typically encountered
with GPCR solubilization and purification. To address this
limitation, a new strategy that uses in vitro GPCR synthesis
in the presence of a previously established,54,55 solidsupported, peptide tethered lipid membrane (tBLM) has been
developed.19 Using an approach similar to that described in
section 3.1, a hydrophilic peptide spacer was covalently
attached to gold slides through amino-terminal thiol groups.
The carboxyl group of the peptide was subsequently activated
and amino-coupled to dimyristoyl-l-R-phosphatidylethanolamine (DMPE), forming a lipid monolayer. The monolayer
was subsequently fused with lipid vesicles to create the final
bilayer. The odorant receptor, OR5 from Rattus norVegicus,
was expressed in vitro directly on the sensor surface.19 This
caused spontaneous integration of the GPCR into the tBLM
upon biosynthesis. The procedure was monitored by surface
plasmon enhanced fluorescence spectroscopy (SPFS) where
fluorescent antibody binding to the GPCR created signals
when the fluorophore was close to the surface. The OR5
receptor was successfully oriented on the surface in such a
way that it mimics the orientation in the native endoplasmic
reticulum, with the N-terminus facing the extracellular side.
Ligand binding was monitored by surface-enhanced infrared
reflection absorption spectroscopy (SERIAS) where absorbance differences were linked to effective binding of lilial,
a small hydrophobic ligand for OR5. Details were not
provided concerning the stability of such slides, and although
this application requires fluorescently labeled ligands, the
success with GPCRs, which tend to be among the most
challenging MPs due to conformational heterogeneity, instability, and low expression yields, suggests that the
procedure has potential to be generally applied to other MPs
and to analytical methods that do not necessitate labeling,
such as SPR.
developed to provide a more stable environment.64 A planarsupported lipid bilayer (PSLB) composed of 1,2-bis[10-2′,4′(hexadienoyloxy)decanoyl]-sn-glycero-3-phosphocholine (bisSorbPC)65 was used for these studies. The bis-SorbPC can
be covalently cross-linked by exposure to UV light, providing
the necessary resistance to washing. Rhodopsin was solubilized in buffer containing OG for these experiments.66,67
Reconstitution of rhodopsin into the PSLBs was carried out
in a plasmon-waveguide resonance (PWR) cell. The PWR
cell was set up in such a way that UV light could be directed
to it from a Mercury lamp, with a band-pass filter to remove
all visible light, which would irreversibly photoactivate
rhodopsin. More than 95% of the bis-SorbPC was polymerized, as monitored by UV absorbance. PWR spectroscopy
was used to characterize rhodopsin immobilized in the crosslinked PSLBs. Within the PSLBs, the opsin isomerization
and G protein activation could be monitored and rates were
found to be similar to those obtained in a fluid dioleoylphosphatidylcholine (DOPC) bilayer. MP immobilization via this
procedure has the advantage of being completed in a few
hours, resulting in a system with intact and functional
immobilized protein. Although no details were given on the
reversibility of this polymerization, it rendered the lipids
resistant to Triton X-100 treatment. Although this method
has great potential for on-surface purification and immobilization of other GPCRs, preserving their functionality
in such a cross-linked environment may not be as easy to
achieve.
5. Covalent Interactions with Protein with
Subsequent Lipid Reconstitution
5.1. Via Carbodiimide Coupling with Lipid
ReconstitutionsGPCR: Rhodopsin
4.4. Via Polymerized Lipid MonomerssGPCR:
Rhodopsin
Rhodopsin has been used extensively to study efficient
immobilization into lipid environments, but in general, lipids
have been shown to be unstable when used for immobilizing
MPs on chips. This is because the hydrophobic interactions
that keep the lipids adsorbed to the glass are not strong
enough to resist eventual desorption after extensive washing.
To address this issue, cross-linked synthetic lipids were
GPCR biosensor studies require high densities of pure and
functional immobilized protein, and this is the limiting factor
with regards to most GPCRs. To address this limitation, a
technique was developed in a flow cell to allow stepwise
addition and removal of mixed detergent and lipid micelles
and protein to create an on-surface lipid bilayer reconstituted
around a solubilized GPCR.10 In principle, large quantities
650 Chemical Reviews, 2011, Vol. 111, No. 2
of unfolded GPCR could be used as input for this approach.
The gold L1 sensor chip was treated to create covalently
linked carboxymethyl-modified dextran polymer with random
glucose moieties substituted by alkyl groups.63 The alkyl
groups hydrophobically bound to mixed micelles of OG
detergent and POPC lipid as they were injected over the
surface to create the initial lipid layer. OG-solubilized
rhodopsin53 was injected and immobilized on the surface by
both the amide chemistry and the strong hydrophobic
interaction between the surface lipid layer and the OG
micelles. The immobilized rhodopsin was immediately
reconstituted in a POPC bilayer by injection of mixed OG
and POPC micelles over the surface. The technique makes
use of the high critical micellar concentration (CMC) of OG
(25 mM). As buffer is injected over the flow cell, OG
monomers detach much quicker than POPC, and as they
detach and wash away, the remaining POPC micelles
spontaneously fuse into a continuous lipid bilayer over the
sensor surface and the immobilized rhodopsin. The functionality of rhodopsin after immobilization was tested by
surface plasmon resonance with light activation and measurement of transducin dissociation from the membrane as
it consumed guanosine 5′-triphosphate (GTP).53 Signaling
was only detected in the sample flow cell containing
rhodopsin-POPC. There was no signal in the absence of
lipids, which proved that their presence68 and correct
reconstitution69 was necessary for the correct function of
rhodopsin.10,70 The ligand-binding capacity of the receptor
was therefore preserved through this immobilization procedure and could be repeatedly measured. This technique
resulted in reasonably high densities of up to 4 ng/mm2 of
immobilized protein, while the use of a polymer hydrogel
layer improved the stability of the immobilized bilayer.
Although protein solubilization and lipid reconstitution were
fast and straightforward via the commercially available
Biacore systems, defining the correct lipid and detergent
stoichiometries for new target proteins will require significant
optimization.71 The technique, however, can be adapted to
different immobilization chemistries, including high-affinity
interactions with antibodies to capture the protein in a defined
orientation72 (see section 7.2).
6. Specific Immobilization of Protein in the Native
Membrane via Linker
Fru¨h et al.
ligand to its binding site. Specific interactions between the
protein or membrane and the surface have been used in place
of random ones for better control of the protein’s orientation.
The most common strategies involve the biotin interaction
with streptavidin or avidin.12,73-75 In one case, a biotinylated
Neurokinin-1 receptor was overexpressed in CHO cell lines
and immobilized through a streptavidin linker to a biotin
covered slide.73 The procedure was very simple, consisting
primarily of treating the quartz slides with biotinylated bovine
serum albumin (BSA), which results in a surface resistant
to salt and detergent washes. The multivalent streptavidin
was then used to bridge the biotinylated GPCR to the
biotinylated BSA for immobilization. The cell lysates
containing overexpressed C-terminally biotinylated Neurokinin-1 receptor were then injected into the chamber directly
to yield a specifically oriented immobilization of the receptor.
Total internal reflection fluorescence (TIRF) was used to
detect binding of streptavidin to the BSA-biotin surface and
to detect binding of the fluorescently labeled substance P
(SP), a Neurokinin-1 receptor agonist, to the oriented
receptor. Binding studies revealed functionality of the
receptor in accordance with previous studies76 but used as
little as 1 attomol of receptor due to the controlled orientation
and high sensitivity of TIRF. This simple immobilization
was the first example of biotinylated MP on quartz surfaces
directly from crude cell lysates without purification, thus
allowing one to study, e.g., mechanisms involved in GPCR
and G protein interactions in the native system. This
technique has the potential to be applied to a wide array of
applications, and if higher densities of proteins are needed
for readouts, on-surface purification and enrichment is
possible because of the high affinity of the immobilization
reagents. It was also suggested that integration of biotinylated
lipids into the plasma membrane of cells prior to homogenization could be an alternative, and in some cases,
chemically oxidized silicon surfaces may be used to avoid
interactions with the lipids.74
7. Specific Immobilization of Detergent
Reconstituted Protein via Linker
7.1. Via N-terminus of Protein on Sepharose
ResinsMP: KcsA and DsbB
6.1. Via Biotinylated Protein Bound to
Streptavidin or AvidinsGPCR: Neurokinin-1
Receptor
Many MPs have specific roles in binding extracellular
ligands. Therefore, when immobilizing such proteins to study
their biological activity, it is desirable to orient them in a
controlled manner so as to maximize the accessibility of the
Fragment-based drug discovery, an approach that consists
of screening small molecules (<300 Da) with promising
bioavailability properties that weakly interact with a pharmaceutical target,77 has gained considerable attention in the
pharma industry. Sensitive techniques, such as crystallography and NMR, are required to detect the weak binding
of fragments to the target. However, many biophysical
techniques require large quantities of stable protein in a pure
form, and that is generally not possible with many MPs.
Although 60% of all drugs target MPs,78 it has not proven
Functional Membrane Protein Immobilization Strategies
possible to apply fragment methods to them. Target immobilized NMR screening (TINS)79 has addressed some of
these limitations, such as the issue of nonspecific binding of
fragments to surfactants or lipids, and shows promise as a
fragment screening method on MPs. In TINS, 1D NMR
spectra of the fragments are simultaneously acquired in the
presence of immobilized reference and target MPs in order
to detect specific binders above the nonspecific level of
fragments interacting with detergents. All proteins in this
case were solubilized in dodecylphosphocholine (DPC)
during the purification procedure, which involved simple
metal affinity chromatography followed by gel filtration in
the presence of the detergent. The immobilization procedure
used Schiff’s base chemistry between primary amines of the
protein and aldehyde groups on the commercially available
sepharose resin. The immobilization efficiencies were reported similar to previous studies on soluble proteins, with
a final concentration equivalent of 100 µM of both the
reference and target proteins on the resin. The functionality
of immobilized DsbB was confirmed by an enzymatic assay,
which indicated that immobilized DsbB retained 90% of its
functionality. The functional immobilization of both KcsA
and DsbB was further demonstrated by detecting binding of
known ligands using TINS.79,80 TINS addresses the protein
demand by reusing a single sample of the target to screen
the entire library, thereby requiring <50 nmol of protein.
Immobilization can be achieved via the N-terminus or a
variety of other chemical strategies, making it potentially
broadly applicable to MPs. OmpA has been noted to retain
conformation in a variety of surfactants,81-85 making it a
useful reference. Although the method enabled fragmentbased drug discovery on MPs, there remained several
limitations. A full screen of 1000 fragments required a week
to complete, and most MPs may not be stable in such
conditions, although, in some cases, immobilization has
proven to improve MP stability. Furthermore, the solubilization processes must still be tailored to each protein.
However, alternative solubilization media (see section 9) such
as Amphipols81 and Nanodiscs,14,86 which appear to be
compatible with TINS, offer the possibility to generalize the
procedure.
Chemical Reviews, 2011, Vol. 111, No. 2 651
(β2AR) was specifically labeled with fluorescein at Cys265,
a conformationally sensitive site. The specific labeling of
only Cys265 by the sulfhydryl-reactive fluorescent probe
fluorescein maleimide was possible because the other cysteines of the protein were either inaccessibly located in the
transmembrane domains or nonreactive because of the
presence of disulfide bonds.87 For exploitation of the
biotin-avidin interaction, two strategies were used in
conjunction with an avidin or streptavidin linker. In the first,
the protein was modified with an N-terminal FLAG epitope
and indirectly linked to the avidin surface through a
biotinylated anti-FLAG antibody. In the second strategy, a
second cysteine residue was biotinylated and linked to the
surface through avidin or streptavidin as in section 5. Liganddependent activation of the β2AR was detected with fluorescence microscopy by monitoring changes in fluorescence
intensity upon ligand binding and receptor translocation. The
β2AR immobilized via the antibody displayed nearly identical
responses to an agonist as the receptor in solution. The β2AR
directly immobilized by the biotin tag yielded less consistent
responses that were significantly smaller than for the receptor
coupled via the antibody. Whether this was an artifact of
the immobilization itself or due to biotinylation of the GPCR
was not clear. This method did not require protein reconstitution into a lipid bilayer and could immobilize protein with
minimal loss due to the high-affinity coupling. Although this
detection method required the GPCR to be labeled with a
fluorophore at a sensitive site, the immobilization strategy
allowed the authors to study conformational changes of the
protein and is compatible with array technologies involving
high-throughput screening of MPs on chips for example,
provided solubilization conditions have been found for a
particular MP.
7.3. Via HIS Tag on Quartz SurfacesMP: Ion
Channel 5-HT3R
7.2. Via Tag and Biotinylated Antibody on
BSA-Biotin SurfacessGPCR: β2-Adrenergic
Receptor
High-affinity interactions have been used successfully to
immobilize GPCRs in their native membrane. However,
many applications require pure protein to provide results with
highsignal-to-noiseratios.Here,thehigh-affinitystreptavidin-biotin
interaction has been used to capture detergent-solubilized
GPCRs in a controlled and oriented manner and monitor
activity by fluorescence microscopy.12 To detect ligand
binding, the detergent-solubilized β2-adrenergic receptor
Many applications require pure protein immobilized at
very high density. Immobilized metal affinity chromatography (IMAC) has been used to successfully immobilize the
detergent-solubilized serotonin-gated ion channel 5-HT3R to
quartz slides without the need for further lipid reconstitution.88 This approach may be generally applicable to all MPs
that can be solubilized and functionally expressed with a HIS
tag. Before immobilization, the glass slides were first
modified to contain thiol groups by gas-phase silanization.
The thiol groups were then covalently linked to a lysine
derivative of nitriloacetic acid (NTA) using a bifunctional
maleimide-succinimide cross-linker. The newly created
surface was fused to a Teflon spacer to create a flow-through
cell and was subsequently charged with Ni2+. The 5-HT3R
ion channel, containing a HIS tag, was solubilized in
nonaethyleneglycol monododecyl and immobilized via chelation to the Ni2+-NTA. The metal affinity-based immobilization procedure was reversible, allowing quantifica-
652 Chemical Reviews, 2011, Vol. 111, No. 2
tion of immobilized receptors by elution with imidazole.
Total internal reflection fluorescence spectroscopy was used
to carry out competition binding experiments using the
nonlabeled competitor quipazine against GR-fluorescein
([1,2,3,9-tetrahydro-3-[(5-methyl-1H-imidazol-4-yl)methyl]9-(3-amino-(N-fluoresceinthiocarbamoyl)propyl)-4H-carbazol-4-one]). The affinity of GR-fluorescein for the receptor
in solution was identical to the surface-bound protein,
demonstrating the feasibility of the process for identifying
potential new drugs and quantifying their affinities with
dose-response curves. This procedure was extremely simple
and could potentially be applied to a variety of analytical
techniques, given the usual constraints that the MP can be
functionally solubilized and expressed with a HIS tag. The
current application is limited to fluorescent ligands, but the
sensitivity of the methods results in signal detection with as
little as 1.6 attomol of immobilized MP, corresponding to
the yield of a single mammalian cell! The detection is realtime, mass-independent, and can be combined with microfluidic applications to further explore high-throughput analysis for drug discovery. Further, given the nature of the
immobilization surface, it seems likely that this procedure
could be readily adapted to SPR applications.
7.4. Via Biotinylated LigandsGPCR: Neurotensin
Receptor-1
SPR analysis of GPCRs is often limited by the fact that
low molecular weight ligands do not generate large signals.
However, by immobilizing the ligand instead, the method
allows one to study GPCR binding to biotinylated ligands
immobilized to streptavidin-covered Biacore chips,89 without
the need for lipid reconstitution. For these studies, the
Neurotensin receptor-1 was expressed in E. coli as an
N-terminal fusion product with the maltose binding protein
and a C-terminal His tag for stability and purification
purposes. Receptor purification by IMAC and neurotensin
affinity chromatography preceded solubilization in detergent
micelles containing (3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate), n-dodecyl-β-maltoside (DDM), and
cholesteryl hemisuccinate (CHS). The peptide ligand Neurotensin (NT) was N-terminally biotinylated and immobilized
onto the streptavidin-coated Biacore chips as the positive
control. A ligand containing a scrambled version of the
primary structure was also synthesized and immobilized on
a second streptavidin-coated chip as a negative control.
Binding was monitored by SPR by flushing the flow cells
with the detergent-solubilized receptor, and highly specific
interactions could be observed and confirmed. The chips
could be regenerated by uncoupling bound receptors with
high salt washes. The authors reported lower amplitude
response than expected at higher levels of immobilized
ligand, which they explained by potential ligand occlusion
that could be prevented by the use of a longer linker between
the surface and the ligand. Although no quantification of the
binding affinities was provided, the technique has potential
Fru¨h et al.
in array technology provided small molecular weight ligands
are available and biotinylation does not affect their binding
affinities to the target protein studied. Further, the technique
could also be adapted to imaging-mode SPR.90 Although this
technique was possible with detergent-solubilized protein,
the limitation of finding appropriate detergents for protein
solubility remains a protein-specific issue.
8. Specific Immobilization of Lipid Reconstituted
Protein via Linker
8.1. Via Biotinylated Proteins in a Mixed
Self-Assembled Monolayer (SAM)sGPCR:
Rhodopsin
To study MPs, which specifically bind ligands from the
extracellular side of the membrane, methods have tended
toward developing controlled, oriented immobilization in
order to expose the appropriate side of the protein to the
ligands. For such a controlled immobilization, a specific
chemistry reacting to the appropriate terminus of the protein
is necessary. Here, carbohydrate-specific chemistry91,92 for
N-terminal biotinylation of glycosylated proteins was combined with thiolipids to anchor rhodopsin to gold surfaces.93
Gold sensor chip surfaces were covered with a homogeneous
self-assembled monolayer (SAM) of ω-hydroxyundecanethiol (HTA), interspaced with biotin attached to the surface
through thiol groups, by micropatterned printing. The HTA
layer was created to avoid protein immobilization in these
areas. Subsequent addition of BSA blocked all nonspecific
binding sites, and addition of streptavidin provided appropriate high-affinity binding sites for the biotinylated receptor.
The glycosylated receptor was specifically biotinylated
on the carbohydrate chains near the N-terminus by oxidizing
the carbohydrate moieties with NaIO4 prior to adding biotin
hydrazide.91 Because only the extracellular facing portions
of the receptor are glycosylated, rhodopsin was immobilized
with the intracellular side facing away from the surface,
allowing for maximal interaction with G proteins. Surface
plasmon resonance in the presence of GTP demonstrated the
possibility of studying the receptor’s constitutive activity.
Interaction with the G protein was directly observed after a
flash of light, and the initial slope of the desorption signal
was a good approximation of receptor density/unit area of
surface. Relaxation of the activated G protein was also
measured, and binding of agonists such as 11-cis- or 9-cisretinal was used to show that the immobilized rhodopsin was
functional. This immobilization strategy stabilized the GPCR
for several hours and multiple cycles of ligand addition and
removal, for which activation could easily be monitored.
Furthermore, due to micropatterns with and without the
receptor, nonspecific binding was calculated as a localized
reference, adding robustness to the data not otherwise
available in SPR analyses. Although this particular biotiny-
Functional Membrane Protein Immobilization Strategies
Chemical Reviews, 2011, Vol. 111, No. 2 653
lation method may be limited to MPs with glycosylated
regions near the targeted terminus, it is an attractive method
for studying G protein mechanisms. However, when applying
MPs in native environments, SPR often proves to be less
sensitive than when used with the well-behaved and highly
overexpressed proteins such as rhodopsin.
properties are more compatible with modern techniques such
as structural biology or biophysical MP-ligand characterization. Amphipols and nanodiscs, which have been used for
MP immobilization, are discussed here, whereas a third,
SMALPs,101 shows promise but has yet to be used for
immobilization.
8.2. Via Lipid Bilayer Tethered through
HIS-Tagged ProteinsMP: Cytochrome C Oxidase
9.1. Via Biotinylated AmphipolsVarious MPs
Studying MPs by electrochemistry is often limited by the
insulating properties of the lipids or detergents if they are
applied directly onto the metallic surface. For detection of
electron transfer, the protein therefore needs to be immobilized away from the surface. Here, a combination of
lipid reconstitution and tethering the target protein through
a HIS tag was used to immobilize cytochrome C oxidase
(CcO) to a silver surface.94,95 The surfaces were roughened
by electrochemical processes and functionalized with Nhydroxysuccinimide ester (NHS) groups by addition of
dithiobis-(N-succinimidyl propionate). An ion-chelating layer
of nitrilotriacetic acid groups (NTA) was finally created by
binding the terminal amino groups of N-(5-amino-1-carboxypentyl) iminodiacetic acid to the existing NHS layer.
Complexation of this new NTA monolayer layer with Cu2+
ions made it possible to reversibly immobilize HIS-tagged
proteins. Immobilization was simply accomplished by flowmediated addition of C-terminally HIS-tagged CcO in DDM.
Finally, the lipid bilayer was established around the receptor
by incubation with a buffer solution of 1,2-diphytanoyl-snglycero-3-phosphocholine (DPGPC) with subsequent removal of DDM. The heme groups of CcO remained intact
and functional upon immobilization, as the ferric and ferrous
states of hemes a and a3 could be monitored repeatedly after
electrode potential changes. The electron transfer rates of
the protein were found to remain unchanged after immobilization, demonstrating an efficient electron exchange
via the HIS tag. An additional benefit of the IMAC
methodology is that it can be used to purify and enrich the
protein in situ.94 This chemistry has been exploited by tagging
C-termini of GPCRs96-99 and is being used to produce highthroughput screening platforms with flow cytometry.100
Details were not supplied regarding stability of the surfaces
or of the immobilized target. Although protein solubilization
remains a challenge, a particularly exciting possibility is the
potential to use crude cell lysates with this approach.94
9. Specific Immobilization of Alternatively
Reconstituted Protein via Linker
Until very recently, only a limited number of options have
been available to solubilize MPs, namely, detergents or
various lipid assemblies. However, three new media have
been proposed as exciting alternatives whose chemical
Amphipols (Apols) are synthetic polymers with a hydrophilic backbone (in this case, polyacrylate) grafted with
various hydrophobic side chains. The amphipathic nature of
Apols enables them to stably bind the transmembrane portion
of MPs to generate a soluble complex. A variety of MPs
have been successfully reconstituted into Apols including
GPCRs. In the present case, the authors reported functional
solubilization of bacteriorhodopsin, the nicotinic acetylcholine receptor, the cytochrome b6f and bc1 complexes, and
the transmembrane domain of E. coli outer membrane protein
A.102 The Apol was chemically modified to include a biotin
moiety prior to reconstitution. Subsequently, streptavidin was
immobilized on one of two surfaces, and the Apol-MP
complex was trapped via noncovalent binding. In one
application, a streptavidin-coated sensor chip was used to
immobilize the Apol-MP complex for SPR studies. Specific
binding of antibodies to each protein was subsequently
observed in a Biacore instrument. Binding of a fluorescent
ligand for the nicotinic acetylcholine receptor, immobilized
on streptavidin-coated beads, was detected by fluorescence
microscopy. A particularly exciting aspect of the work is
the general usefulness of Apols for functional solubilization
of MPs, a point that does not escape the author’s attention.
9.2. Via the N-terminus of Nanodisc Solubilized
Protein MP: KcsA and DsbB
Nanodiscs (NDs) are lipid bilayers in which the exposed
hydrophobic side chains at the edge of the disk-shaped
complex are protected by binding of an amphipathic,
R-helical protein. Nanodiscs are advantageous in that they
are an excellent membrane mimetic and yet have a welldefined and relatively homogeneous structure. Recently, the
bacterial proteins KcsA and DsbB were functionally reconstituted into NDs and immobilized via Schiff’s base chemistry.83 In this case, the immobilization was partially via the
654 Chemical Reviews, 2011, Vol. 111, No. 2
MP and partly via the ND, but in principle, an appropriate
affinity tag could be placed or engineered on the ND such
that immobilization would occur exclusively via the ND. The
ND-solubilized protein was compared to micelle-solubilized
protein for biophysical ligand-screening studies. It was shown
that there is significantly more nonspecific binding of small
molecules to micelles than to NDs and that this nonspecific
binding occurs more frequently with more hydrophobic small
molecules. The nonspecific binding can mask specific
binding, thereby hindering the ability to assess whether a
large portion of the library binds to the MP of interest.
10. Perspectives
Biophysical assays of protein function and/or ligand
binding are playing an ever-increasing role in both academic
and industrial life science research. Applications in academic
laboratories include determining and understanding protein
interaction networks that lead to regulation of cell behavior
(e.g., proteomics) while the primary use in industry has been
for the discovery of ligands that modulate protein behavior.
To scale these assays down and/or make efficient use of
limited resources, the proteins are often immobilized on flat
metallic or glass surfaces with little or no biocompatible
characteristics. Although the array of biophysical techniques
that have been successfully applied to soluble proteins is
impressive, until recently similar applications to membranebound proteins were sparse. Here we have highlighted many
of the success stories culled from the literature of the past
few years. One common denominator of these success stories
is the innovation and effort required to overcome the
bioincompatibility of the surfaces. A second recurring theme
is that each solution must be tailored to the individual protein
being studied. Together these two remaining issues represent
a bottleneck to widespread, high-throughput biophysical
assays that could take advantage of, for example, printing
techniques.
We see two developments that, when combined with
methodologies that have been described, offer exciting
opportunities for more generic application of biophysical
techniques. Interestingly, these developments come from both
sides of the problem, that is, the protein itself and the media
used to solubilize the protein, and therefore are potentially
complementary. Approaching the problem from the protein
point of view, the group of Christopher Tate at Cambridge
University (U.K.) has developed a technique for selecting
mutants that provide enhanced thermal and conformational
stability while retaining desired ligand-binding properties.103
Importantly, these stabilized proteins are compatible with a
much wider range of detergents than the wild-type protein
and thus should be more amenable to large-scale immobilization studies. From the media point of view, important advances have been made in developing alternatives to
detergentsforfunctionalsolubilizationofMPs.Nanodiscs,14,86,104
bilayer lipid assemblies surrounded by a stabilizing protein,
and amphipols,105-107 synthetic amphipathic polymers, have
successfully replaced detergents to solubilize a variety of
MPs including GPCRs. Lipidic sponge phases represent an
even more recent addition to this list, which, so far, shows
promise for crystallization of MPs.108 The crucial advantage
of these new media is that they represent a more or less “one
size fits all” solution that holds the promise of eliminating,
or at least greatly minimizing, the requirement to precisely
tailor each solution to the MP.
Fru¨h et al.
Ultimately, applications that combine the high sensitivity
of biophysical methods with the possibility of studying an
MP in its native environment without the need for purification
and reconstitution represent the Holy Grail for many research
goals. While at present still out of reach, the growing body
of information on production and immobilization strategies
and the ever-increasing sophistication of biophysical methods
will undoubtedly conspire to enable this in situ approach.
The combination of such biophysical studies with increasing
success in the crystallization and NMR analysis of MPs
should prove a powerful approach to both reveal molecular
mechanisms of membrane protein function and enable
rational elaboration of small-molecule modulators of this
function.
11. Abbreviations
5-HT3
Apol
BSA
DsbB
CB
CcO
C8E4
CHAPS
CHS
DDM
DMPC
DMPE
DPC
DPGPC
DPPA
DPPC
DPPE
FRAP
GAPS
GR
HTA
IMAC
KcsA
LB
nAChR
ND
NMP
NMR
NTA
OG
OmpA
OTG
PC
POPC
POPG
PSLB
PWR
SAM
SERIAS
SPA
SPFS
SPR
tBLM
TIFR
serotonin-gated mouse ion channel 5-hydroxytryptamine type-3 receptor
Amphipol
bovine serum albumin
disulfide bond-forming protein B
cytochalasin B
cytochrome C oxidase
n-octyltetraoxyethylene
3-[(3-cholamidopropyl)dimethylammonio]-1propanesulfonate
cholesteryl hemisuccinate
n-dodecyl-β-maltoside
dimyristylphosphatidylcholine
1,2-dimyristoyl-sn-glycerophosphatidylethanolamine
dodecylphosphocholine
1,2-diphytanoyl-sn-glycero-3-phosphocholine
dipalmitoyl L-R-phosphatidic acid
dipalmitoyl phosphatidylcholine
1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine
fluorescence recovery after photobleaching
γ-aminopropylsilane
1,2,3,9-tetrahydro-3-[(5-methyl-1H-imidazol-4yl)methyl]-9-[3-amino-(N-fluoresceinthiocarbamoyl)propyl]-4H-carbazol-4-one
ω-hydroxyundecanethiol
immobilized metal affinity chromatography
K+ channel from Streptomyces liVidans
Langmuir-Blodgett
nicotinic acetylcholine receptor
nanodisc
N-methylpyrolidone
nuclear magnetic resonance
nitrilotriacetic acid
octylglucoside
outer membrane protein A
n-octyl-D-thioglucopyranoside
phosphatidylcholine
palmitoyl oleoyl phosphatidylcholine
palmitoyl oleoyl phosphatidylglycerol
planar-supported lipid bilayers
plasmon-waveguide resonance
self-assembled monolayer
surface-enhanced infrared reflection absorption spectroscopy
scintillation proximity assay
surface plasmon enhanced fluorescence spectroscopy
surface plasmon resonance
tethered bilayer lipid membrane
total internal reflection fluorescence microscopy
Functional Membrane Protein Immobilization Strategies
12. Acknowledgments
The authors thank Prof. Karen Martinez for carefully
reading the manuscript.
13. References
(1) Marassi, F. M.; Ramamoorthy, A.; Opella, S. J. Proc. Natl. Acad.
Sci. U. S. A. 1997, 94, 8551.
(2) Leiefert, W. R.; Aloia, A. L.; Bucco, O.; Glatz, R. V.; McMurchie,
E. J. J. Biomol. Screening 2005, 10, 765.
(3) MacBeath, G.; Schreiber, S. L. Science 2000, 289, 1760.
(4) Bertucci, C.; Cimitan, S. J. Pharm. Biomed. Anal. 2003, 32, 707.
(5) Arenkov, P.; Kukhtin, A.; Gemmell, A.; Voloshchuk, S.; Chupeeva,
V.; Mirzabekov, A. Anal. Biochem. 2000, 278, 123.
(6) Guschin, D.; Yershov, G.; Zaslavsky, A.; Gemmell, A.; Shick, V.;
Proudnikov, D.; Arenkov, P.; Mirzabekov, A. Anal. Biochem. 1997,
250, 203.
(7) Lu, L.; Brekkan, E.; Haneskog, L.; Yang, Q.; Lundahl, P. Biochim.
Biophys. Acta 1993, 1150, 135.
(8) Yang, Q.; Lundahl, P. Anal. Biochem. 1994, 218, 210.
(9) Gottschalk, I.; Li, Y. M.; Lundahl, P. J. Chromatogr., B 2000, 739,
55.
(10) Karlsson, O. P.; Lofas, S. Anal. Biochem. 2002, 300, 132.
(11) Franzin, C. M.; Gong, X. M.; Thai, K.; Yu, J. H.; Marassi, F. M.
Methods 2007, 41, 398.
(12) Neumann, L.; Wohland, T.; Whelan, R. J.; Zare, R. N.; Kobilka,
B. K. ChemBioChem 2002, 3, 993.
(13) Naumann, R.; Jonczyk, A.; Hampel, C.; Ringsdorf, H.; Knoll, W.;
Bunjes, N.; Graber, P. Bioelectrochem. Bioenergy 1997, 42, 241.
(14) Civjan, N. R.; Bayburt, T. H.; Schuler, M. A.; Sligar, S. G.
BioTechniques 2003, 35, 556.
(15) Gottschalk, I.; Lagerquist, C.; Zuo, S. S.; Lundqvist, A.; Lundahl,
P. J. Chromatogr., B 2002, 768, 31.
(16) Schmid, E. L.; Tairi, A. P.; Hovius, R.; Vogel, H. Anal. Chem. 1998,
70, 1331.
(17) Fang, Y.; Frutos, A. G.; Lahiri, J. ChemBioChem 2002, 3, 987.
(18) Vanwetswinkel, S.; Heetebrij, R. J.; van Duynhoven, J.; Hollander,
J. G.; Filippov, D. V.; Hajduk, P. J.; Siegal, G. Chem. Biol 2005,
12, 207.
(19) Robelek, R.; Lemker, E. S.; Wiltschi, B.; Kirste, V.; Naumann, R.;
Oesterhelt, D.; Sinner, E. K. Angew. Chem., Int. Ed. 2007, 46, 605.
(20) Adair, W. L.; Kornfeld, S. J. Biol. Chem. 1974, 249, 4696.
(21) Moaddel, R.; Oliveira, R. V.; Hyppolite, P.; Juhaszova, M.; Wainer,
I. W. J. Pharm. Biomed. Anal. 2010, 52, 416.
(22) Perez, J. B.; Martinez, K. L.; Segura, J. M.; Vogel, H. AdV. Funct.
Mater. 2006, 16, 306.
(23) Perez, J. B.; Segura, J. M.; Abankwa, D.; Piguet, J.; Martinez, K. L.;
Vogel, H. J. Mol. Biol. 2006, 363, 918.
(24) Danelon, C.; Perez, J. B.; Santschi, C.; Brugger, J.; Vogel, H.
Langmuir 2006, 22, 22.
(25) Pick, H.; Schmid, E. L.; Tairi, A. P.; Ilegems, E.; Hovius, R.; Vogel,
H. J. Am. Chem. Soc. 2005, 127, 2908.
(26) Bayley, H.; Cremer, P. S. Nature 2001, 413, 226.
(27) Stamou, D.; Duschl, C.; Delamarche, E.; Vogel, H. Angew. Chem.,
Int. Ed. Engl. 2003, 42, 5580.
(28) Hart, H. E.; Greenwald, E. B. Mol. Immunol 1979, 16, 265.
(29) Gruner, S. M.; Kirk, G.; Patel, L.; Kaback, H. R. Biochemistry 1982,
21, 3239.
(30) Nelson, N. Anal. Biochem. 1987, 165, 287.
(31) Bosse, R.; Garlick, R.; Brown, B.; Menard, L. J. Biomol. Screen.
1998, 3, 285.
(32) Simpson, P. R.; Yu, X. H.; Redza, Z. M.; Anson, J. G.; Chan, S. H.;
Lin, Y. J. Virol. Methods 1997, 69, 197.
(33) Gevi, M.; Domenici, E. Anal. Biochem. 2002, 300, 34.
(34) Lerner, C. G.; Saiki, A. Y. C. Anal. Biochem. 1996, 240, 185.
(35) Trivedi, V.; Zhang, S. C.; Castoreno, A. B.; Stockinger, W.; Shieh,
E. C.; Vyas, J. M.; Frickel, E. M.; Nohturfft, A. Proc. Natl. Acad.
Sci. U. S. A. 2006, 103, 18226.
(36) Lu, L.; Brekkan, E.; Haneskog, L.; Yang, Q.; Lundahl, P. Biochim.
Biophys. Acta 1993, 1150, 135.
(37) Yang, Q.; Lundahl, P. Anal. Biochem. 1994, 218, 210.
(38) Sandberg, M.; Lundahl, P.; Greijer, E.; Belew, M. Biochim. Biophys.
Acta 1987, 924, 185.
(39) Kasahara, M.; Hinkle, P. C. J. Biol. Chem. 1977, 252, 7384.
(40) Kirby, C.; Gregoriadis, G. Biotechnol. J. 1984, 2, 979.
(41) Brekkan, E.; Lundqvist, A.; Lundahl, P. Biochemistry 1996, 35,
12141.
(42) Bayer, R.; Feigenson, G. W. Biochim. Biophys. Acta 1985, 815, 369.
(43) Puu, G.; Koch, M. Biochem. Pharmacol. 1990, 40, 2209.
(44) Sellstrom, A.; Gustafson, I.; Ohlsson, P. A.; Olofsson, G.; Puu, G.
Colloids Surf. 1992, 64, 289.
Chemical Reviews, 2011, Vol. 111, No. 2 655
(45) Puu, G.; Gustafson, I.; Artursson, E.; Ohlsson, P. A. Biosens.
Bioelectron. 1995, 10, 463.
(46) Posner, B.; Hong, Y. L.; Benvenuti, E.; Potchoiba, M.; Nettleton,
D.; Lui, L.; Ferrie, A.; Lai, F.; Fang, Y.; Miret, J.; Wielis, C.; Webb,
B. Anal. Biochem. 2007, 365, 266.
(47) Fang, Y.; Lahiri, J.; Picard, L. Drug DiscoVery Today 2003, 8, 755.
(48) Fang, Y.; Frutos, A. G.; Lahiri, J. ChemBioChem 2002, 3, 987.
(49) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124,
2394.
(50) Lahiri, J.; Kalal, P.; Frutos, A. G.; Jonas, S. T.; Schaeffler, R.
Langmuir 2000, 16, 7805.
(51) Carre, A. R. E.; Efrenov, A. M.; Fang, Y.; Hong, Y.; Lacarriere, V.;
Lahiri, J.; Lai, F.; Mauro, J. C.; Raghavan, S.; Webb, B. L. Patent
20060147993 2006.
(52) Bailey, K.; Bally, M.; Leifert, W.; Vo¨ro¨s, J.; McMurchie, T.
Proteomics 2009, 9, 2052.
(53) Heyse, S.; Ernst, O. P.; Dienes, Z.; Hofmann, K. P.; Vogel, H.
Biochemistry 1998, 37, 507.
(54) Wiltschi, B.; Schober, M.; Kohlwein, S. D.; Oesterhelt, D.; Sinner,
E. K. Anal. Chem. 2006, 78, 547.
(55) Sinner, E. K.; Reuning, U.; Kok, F. N.; Sacca, B.; Moroder, L.; Knoll,
W.; Oesterhelt, D. Anal. Biochem. 2004, 333, 216.
(56) Naumann, R.; Schmidt, E. K.; Jonczyk, A.; Fendler, K.; Kadenbach,
B.; Liebermann, T.; Offenhausser, A.; Knoll, W. Biosens. Bioelectron.
1999, 14, 651.
(57) Bunjes, N.; Schmidt, E. K.; Jonczyk, A.; Rippmann, F.; Beyer, D.;
Ringsdorf, H.; Graber, P.; Knoll, W.; Naumann, R. Langmuir 1997,
13, 6188.
(58) Schmidt, G.; Graber, P. Biochim. Biophys. Acta 1985, 808, 46.
(59) Boekema, E. J.; Schmidt, G.; Graber, P.; Berden, J. A. Z. Naturforsch.,
C 1988, 43, 219.
(60) Kadenbach, B.; Stroh, A.; Ungibauer, M.; Kuhnnentwig, L.; Buge,
U.; Jarausch, J. Method Enzymol. 1986, 126, 32.
(61) Minic, J.; Grosclaude, J.; Aioun, J.; Persuy, M. A.; Gorojankina, T.;
Salesse, R.; Pajot-Augy, E.; Hou, Y. X.; Helali, S.; Jaffrezic-Renault,
N.; Bessueille, F.; Errachid, A.; Gomila, G.; Ruiz, O.; Samitier, J.
Biochim. Biophys. Acta 2005, 1724, 324.
(62) Vidic, J.; Pla-Roca, M.; Grosclaude, J.; Persuy, M. A.; Monnerie,
R.; Caballero, D.; Errachid, A.; Hou, Y. X.; Jaffrezic-Renault, N.;
Salesse, R.; Pajot-Augy, E.; Samitier, J. Anal. Chem. 2007, 79, 3280.
(63) Cooper, M. A.; Hansson, A.; Lofas, S.; Williams, D. H. Anal.
Biochem. 2000, 277, 196.
(64) Subramaniam, V.; Alves, I. D.; Salgado, G. F. J.; Lau, P. W.;
Wysocki, R. J.; Salamon, Z.; Tollin, G.; Hruby, V. J.; Brown, M. F.;
Saavedra, S. S. J. Am. Chem. Soc. 2005, 127, 5320.
(65) Sisson, T. M.; Lamparski, H. G.; Kolchens, S.; Elayadi, A.; O’Brien,
D. F. Macromolecules 1996, 29, 8321.
(66) Botelho, A. V.; Gibson, N. J.; Thurmond, R. L.; Wang, Y.; Brown,
M. F. Biochemistry 2002, 41, 6354.
(67) Alves, I. D.; Salamon, Z.; Varga, E.; Yamamura, H. I.; Tollin, G.;
Hruby, V. J. J. Biol. Chem. 2003, 278, 48890.
(68) Brown, M. F. Chem. Phys. Lett. 1994, 73, 159.
(69) Bubis, J. Biol. Res 1998, 31, 59.
(70) Schmitt, J.; Danner, B.; Bayerl, T. M. Langmuir 2001, 17, 244.
(71) Schurholz, T. Biol. Psychiatry 1996, 58, 87.
(72) Stenlund, P.; Babcock, G. J.; Sodroski, J.; Myszka, D. G. Anal.
Biochem. 2003, 316, 243.
(73) Martinez, K. L.; Meyer, B. H.; Hovius, R.; Lundstrom, K.; Vogel,
H. Langmuir 2003, 19, 10925.
(74) Lei, S. B.; Tero, R.; Misawa, N.; Yamamura, S.; Wan, L. J.; Urisu,
T. Chem. Phys. Lett. 2006, 429, 244.
(75) Kada, G.; Riener, C. K.; Hinterdorfer, P.; Kienberger, F.; Stoh, C. M.;
Gruber, H. J. Single Mol. 2002, 3, 119.
(76) Tota, M. R.; Daniel, S.; Sirotina, A.; Mazina, K. E.; Fong, T. M.;
Longmore, J.; Strader, C. D. Biochemistry 1994, 33, 13079.
(77) Hann, M. M.; Leach, A. R.; Harper, G. J. Chem. Inf. Comp. Sci.
2001, 41, 856.
(78) Zheng, C. J.; Han, L. Y.; Yap, C. W.; Xie, B.; Chen, Y. Z. Drug
DiscoVery Today 2006, 11, 412.
(79) Fru¨h, V.; Heetebrij, R.; Siegal, G. Fragment-Based Drug DiscoVery:
A Practical Approach; John Wiley & Sons: Chichester, U.K., 2008;
Chapter Ch. 6, p 135.
(80) Fru¨h, V.; Zhou, Y.; Chen, D.; Loch, C.; AB, E.; Grinkova, Y. N.;
Verheij, H.; Sligar, S. G.; Bushweller, J. H.; Siegal, G. Chem. &
Bio. 2010, 17, 881.
(81) Zoonens, M.; Giusti, F.; Zito, F.; Popot, J. L. Biochemistry 2007,
46, 10392.
(82) Bond, P. J.; Sansom, M. S. P. J. Mol. Biol. 2003, 329, 1035.
(83) Arora, A.; Abildgaard, F.; Bushweller, J. H.; Tamm, L. K. Nat. Struct.
Biol. 2001, 8, 334.
(84) Pautsch, A.; Schulz, G. E. Nat. Struct. Biol. 1998, 5, 1013.
(85) Pautsch, A.; Schulz, G. E. J. Mol. Biol. 2000, 298, 273.
656 Chemical Reviews, 2011, Vol. 111, No. 2
(86) Bayburt, T. H.; Carlson, J. W.; Sligar, S. G. J. Struct. Biol 1998,
123, 37.
(87) Ghanouni, P.; Steenhuis, J. J.; Farrens, D. L.; Kobilka, B. K. Proc.
Natl. Acad. Sci. U.S.A. 2001, 98, 5997.
(88) Schmid, E. L.; Tairi, A. P.; Hovius, R.; Vogel, H. Anal. Chem. 1998,
70, 1331.
(89) Harding, P. J.; Hadingham, T. C.; McDonnell, J. M.; Watts, A. Eur.
Biophys. J. Biophys. 2006, 35, 709.
(90) Neumann, T.; Junker, H. D.; Schmidt, K.; Sekul, R. Curr. Top. Med.
Chem. 2007, 7, 1630.
(91) Oshannessy, D. J.; Dobersen, M. J.; Quarles, R. H. Immunol. Lett.
1984, 8, 273.
(92) Spinke, J.; Liley, M.; Schmitt, F. J.; Guder, H. J.; Angermaier, L.;
Knoll, W. J. Chem. Phys. 1993, 99, 7012.
(93) Bieri, C.; Ernst, O. P.; Heyse, S.; Hofmann, K. P.; Vogel, H. Nat.
Biotechnol. 1999, 17, 1105.
(94) Friedrich, M. G.; Giess, F.; Naumann, R.; Knoll, W.; Ataka, K.;
Heberle, J.; Hrabakova, J.; Murgida, D. H.; Hildebrandt, P. Chem.
Commun. 2004, 2376.
(95) Giess, F.; Friedrich, M. G.; Heberle, J.; Naumann, R. L.; Knoll, W.
Biophys. J. 2004, 87, 3213.
(96) Sklar, L. A.; Vilven, J.; Lynam, E.; Neldon, D.; Bennett, T. A.;
Prossnitz, E. BioTechniques 2000, 28, 976.
(97) Sklar, L. A.; Simons, P.; Waller, A.; Biggs, S.; Buranda, T.; Foutz,
T.; Edwards, B. S.; Prossnitz, E. R. Cytometry, Part A 2004, 59A,
94.
Fru¨h et al.
(98) Waller, A.; Simons, P.; Prossnitz, E. R.; Edwards, B. S.; Sklar, L. A.
Comb. Chem. High Throughput Screening 2003, 6, 389.
(99) Waller, A.; Simons, P. C.; Biggs, S. M.; Edwards, B. S.; Prossnitz,
E. R.; Sklar, L. A. Trends Pharmacol. Sci. 2004, 25, 663.
(100) Sklar, L. A.; Carter, M. B.; Edwards, B. S. Curr. Opin. Pharmacol
2007, 7, 527.
(101) Knowles, P. F.; Finka, R.; Smith, C.; Lin, Y. P.; Dafforn, T.;
Overduin, M. J. Am. Chem. Soc. 2009, 131, 7484.
(102) Charvolin, D.; Perez, J.-B.; Rouvie`re, F.; Giusti, F.; Bazzacco, P.;
Abdine, A.; Martinez, K. L.; Popot, J.-L. Proc. Natl. Acad. Sci. U.S.A.
2009, 106, 405.
(103) Serrano-Vega, M. J.; Magnani, F.; Shibata, Y.; Tate, C. G. Proc.
Natl. Acad. Sci. U.S.A. 2008, 105, 877.
(104) Leitz, A. J.; Bayburt, T. H.; Barnakov, A. N.; Springer, B. A.; Sligar,
S. G. BioTechniques 2006, 40, 601.
(105) Champeil, P.; Menguy, T.; Tribet, C.; Popot, J. L.; Le Maire, M.
J. Biol. Chem. 2000, 275, 18623.
(106) Zoonens, M.; Catoire, L. J.; Giusti, F.; Popot, J. L. Proc. Natl. Acad.
Sci. U. S. A. 2005, 102, 8893.
(107) Diab, C.; Tribet, C.; Gohon, Y.; Popot, J. L.; Winnik, F. M. Biochim.
Biophys. Acta, Biomembr. 2007, 1768, 2737.
(108) Wohri, A. B.; Johansson, L. C.; Wadsten-Hindrichsen, P.; Wahlgren,
W. Y.; Fischer, G.; Horsefield, R.; Katona, G.; Nyblom, M.; Oberg,
F.; Young, G.; Cogdell, R. J.; Fraser, N. J.; Engstrom, S.; Neutze,
R. Structure 2008, 16, 1003.
CR900088S